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Control of Heme Coordination and Catalytic Activity by Conformational Changes in Peptide−Amphiphile Assemblies Lee A. Solomon,† Jacob B. Kronenberg,‡ and H. Christopher Fry*,† †

Argonne National Laboratory, 9700 South Cass Avenue, Argonne, Illinois 60439, United States Illinois Math and Science Academy, 1500 West Sullivan Road, Aurora, Illinois 60506, United States

‡

S Supporting Information *

ABSTRACT: Self-assembling peptide materials have gained significant attention, due to well-demonstrated applications, but they are functionally underutilized. To advance their utility, we use noncovalent interactions to incorporate the biological cofactor heme-B for catalysis. Heme-proteins achieve differing functions through structural and coordinative variations. Here, we replicate this phenomenon by highlighting changes in heme reactivity as a function of coordination, sequence, and morphology (micelles versus fibers) in a series of simple peptide amphiphiles with the sequence c16-xyL3K3-CO2H where c16 is a palmitoyl moiety and xy represents the heme binding region: AA, AH, HH, and MH. The morphology of this peptide series is characterized using transmission electron and atomic force microscopies as well as dynamic light scattering. Within this small library of peptide constructs, we show that three spectroscopically (UV/visible and electron paramagnetic resonance) distinct heme environments were generated: noncoordinated/embedded high-spin, five-coordinate high-spin, and six-coordinate low-spin. The resulting material’s functional dependence on sequence and supramolecular morphology is highlighted 2-fold. First, the heme active site binds carbon monoxide in both micelles and fibers, demonstrating that the heme active site in both morphologies is accessible to small molecules for catalysis. Second, peroxidase activity was observed in heme-containing micelles yet was significantly reduced in heme-containing fibers. We briefly discuss the implications these findings have in the production of functional, self-assembling peptide materials.

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INTRODUCTION Controlling catalytic activity through the organized assembly of molecules is at the heart of many biological processes and remains a challenge in the field of de novo protein design and nanoscale architectures.1−9 Natural proteins achieve this through a chain of amino acids that fold into a threedimensional structure that in turn governs molecular organization and activity. Achieving complex biologically relevant reactivity in synthetic materials remains a challenge because of the need to simultaneously balance properties that lead to supramolecular assembly while maintaining precise molecular control in the catalytic active site. Peptide amphiphiles (PAs), a class of supramolecular biomaterials, provide a simple solution to this problem.10−12 The peptide serves as a scaffold that typically includes a recognition, structural, and functional site. The structural region guides assembly, while the functional site, typically a modified or unnatural amino acid or sequence of amino acids, is used in catalysis. 2,13−23 The peptide, however, can be further programmed to incorporate added functions, ultimately generating a protein-like catalytic material. In our efforts to design biologically inspired materials, we have engineered metal binding sites into peptide amphiphiles to generate functional supramolecular assemblies.24,25 Transition metal binding peptides, heme binding amyloid-β aggregates, and hemoprotein assemblies have demonstrated catalytic properties.5,26−28 In this © 2017 American Chemical Society

work, we incorporate the naturally occurring cofactor heme-B, inspired by the diversity of heme enzymes, to elevate peptide amphiphiles to a new level of sophistication. We demonstrate a system where the supramolecular structure alone can control heme coordination and reactivity. As a result, we present a peptide-based self-assembling material that functions like a natural protein. Heme-B (Fe-protoporphyrin IX) has an impressively diverse functional library in nature. Uncoordinated, it is toxic due to its ability to produce reactive oxygen species.29,30 However, when associated with a protein, function can be focused toward important metabolic activities. This is due to the heme’s immediate coordination environment, which is significantly influenced by the protein structure.31,32 For example, nitrophorin coordinates heme with a single histidine and, due to its structure, functions as a nitric oxide carrying protein found in insects, whereas neuroglobin uses a bis-histidine coordination to bind oxygen in the brain, Figure 1.33−36 Cytochrome c employs one histidine and one methionine ligand to carry out high potential electron transfer.37−41 At present, no artificial material is able to associate with a single cofactor and carry out such diverse array of functions, but achieving this level of control would add new dimensions to material applications. Received: February 17, 2017 Published: May 15, 2017 8497

DOI: 10.1021/jacs.7b01588 J. Am. Chem. Soc. 2017, 139, 8497−8507

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are well understood in natural systems, but translating them to materials and other technologies has proven difficult.42−44 In this work, we not only demonstrate these long-range conformational changes (i.e., micelle to fiber transition) in peptide−amphiphile assemblies, but also use them to control reactivity of our noncovalently bound cofactor. First, we highlight our ability to vary the heme-binding site through a single “mutation” within the primary sequence that changes the coordination environment around the heme, Figure 1. Second, we present striking evidence that the supramolecular assembly, micelles versus fibers, significantly influences the heme coordination mode. Third, we highlight the material’s ability to bind carbon monoxide, which serves as a redox inactive surrogate to other biologically relevant gases like O2 and NO and thus confirms heme active site accessibility for catalysis. Finally and most intriguingly, we demonstrate our ability to dramatically reduce peroxidase activity solely on the basis of the peptide amphiphile supramolecular structure, micelles versus fibers. These discoveries emphasize the robustness of the peptide amphiphile in developing next generation, functional, biomolecular materials.

Figure 1. Protein structure, active site, and design inspiration for functional heme peptide amphiphiles. Cartoon depictions of crystal structures for nitrophorin (PDB ID 1ERX), neuroglobin (PDB ID 2VRY), and cytochrome c (PDB ID 3CYT). Details of the primary coordination sphere highlighting no-coordination, single histidine, bishistidine, and histidine-methionine. Idealized binding of the designed heme-binding peptide amphiphiles in a β-sheet conformation with their abbreviated names and sequence. Color coding for PA molecules: gray, palmitoyl/c16; yellow, alanine; orange, methionine; red, histidine; green, leucine; and blue, lysine.

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RESULTS AND DISCUSSION Design. The peptide−amphiphiles were designed to emulate naturally occurring heme active sites by reproducing the coordination environments shown in Figure 1. The peptides follow the simple design, c16-xyL3K3-CO2H. We have employed our rational design strategy from our previous study where c16 is a palmitoyl moiety that was included to promote hydrophobic collapse, the first step in peptide amphiphile assembly. The positively charged trilysine (K3) headgroup is introduced as a pH trigger to induce fiber assembly. In other words, raising the pH close to the pKa of the lysine residues reduces electrostatic repulsion, allowing individual peptide molecules to come within van der Waals distances. Three leucine amino acids (L3) serve as a β-sheet structural motif that assists in the formation of long-aspect ratio nanofibers. The heme binding site is represented by x and y, where y is typically histidine (H), the most common hemebinding amino acid in nature, and x represents the site employed to vary the coordination state: A, M, or H (Figure 1). As a result, the binding site xy is generated (AA, AH, HH, or

In addition to coordination, neuroglobin, cytochrome c, and hemoglobin further control heme-function through large-scale conformational changes.42−44 For example, cytochrome c in its native state is an electron transfer protein, but it is also overexpressed in cancerous cells where a conformational change leads to a functional change from an electron transfer protein to a peroxidase. Korendovych et al. were able to redesign a natural protein with existing conformational changes to engineer a switchable eliminase.45 Similarly, Grosset et al. engineered allosteric rearrangements in a de novo protein, using heme as a redox-switch, but were unable to couple that to a function.46 The triggers and effects of long-range interactions

Figure 2. Characterization of supramolecular morphologies in different aqueous solutions. Atomic force micrographs (2 μm × 2 μm) of HH assemblies in HEPES (A) without and (B) with hemin and in 10 mM NH4OH (C) without and (D) with hemin. The height profiles to the right of the micrographs are measurements of individual micelles or fibers without (black) and with hemin (red). 8498

DOI: 10.1021/jacs.7b01588 J. Am. Chem. Soc. 2017, 139, 8497−8507

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drive the micelle to fiber transition (Figure S5). AFM highlights that when heme is coordinated/embedded in the peptide fibers (10:1 peptide:heme), all xyHeme peptide assemblies have the same height profile, 7−8 nm (Figure S3). A relationship between morphology and secondary structure is also noted. In HEPES buffer where micelles are observed, FT-IR analysis indicates that AH and HH lack a well-defined secondary structure as observed by the broad vibrational modes. MH exhibits a β-sheet component with vibrational modes at 1630 and 1681 cm−1. The presence of β-sheets in MH is somewhat anomalous as β-sheets are often linked to fiber formation. This relationship between β-sheets and fibers is observed in AA in HEPES buffer where a high degree of βsheet content is observed with an intense amide I vibration at 1630 cm−1 and a less intense band at 1681 cm−1, Figure S6a.58 We then added heme (at a ratio of 10 peptides:1 heme to ensure binding) to see if it has an effect on structure. Upon addition of heme, no significant change in secondary structure was observed for any of the peptide assemblies (Figure S6b) consistent with the lack of change in observed morphology. At pH 10.5, we detect the formation of β-sheets by FTIR and CD (Figures S6C and S7, respectively) with or without heme (10 peptide:1 heme, Figure S5D). The pKa of lysine is close to 10.5 and becomes neutralized, allowing neighboring molecules to interact and form β-sheets. The FTIR indicates that all xy assemblies, both apo and holo, at higher pH yield amide I vibrations (1630 and 1681 cm−1) consistent with β-sheets. We note that the CD spectra do not yield signature β-sheet spectra with a minimum at 218 nm. The red-shifted spectra (λmin = 218−229 nm) (Figure S7) are attributed to a superhelical twist within the fiber construct, and the observed variations in superhelicity (i.e., degree of red shifting) are influenced by changes in the peptide sequence.59 Therefore, in the xy peptide series, a lack of well-defined secondary structure at neutral pH in HEPES buffer directly correlates to spherical micelle structures, whereas β-sheet formation at high pH directly relates to long aspect ratio fibers in bundled networks. In both micelles and fibers, the overall structure is largely unaffected by the addition of heme. As a result, this peptide series allows the examination of how both the primary amino acid sequence of the peptides and the supramolecular assembly influence the heme cofactor binding and function. Ferric Heme Binding. To investigate how PA sequence and morphology influence heme coordination environment, we employed electronic absorption (EA) spectroscopy and electron paramagnetic resonance (EPR) spectroscopy that together help describe the mechanism of heme insertion, the ligand environment, and the spin state of the metal-centered cofactor, Figure 4 and Table 1. Heme Coordinated to Micelle Characterization (HEPES, pH 7). For spectroscopic characterization, the peptide:heme stoichiometry was maintained at 10:1 to ensure complete heme coordination. In HHHeme micelles at neutral pH, the electronic absorption spectrum yields signature Soret (λmax = 413 nm) and Q-band (λmax = 535, 560 nm) values consistent with bis-histidine axial coordination to heme, in agreement with the spectrum for many bis-histidine coordinating proteins including neuroglobin (Figure 4A, Table 1).60 The EPR spectrum represents a purely S = 1/2, low-spin, type II (rhombic) spin configuration (Figure 4C, Table 1) and is characteristic of many low-spin, bis-histidine coordinated heme proteins including neuroglobin.61−64 AAHeme (fibers at neutral pH) does not offer a histidine containing coordination site and

MH) and is further denoted as xy (e.g., AH) in the apo state (uncoordinated heme) or xyHeme (e.g., AHHeme) in the holo state (coordinated heme). For the initial design visualization, we assumed the formation of parallel β-sheets, typical for amphiphiles (Figure 1). AHHeme was designed to have axial ligation similar to that of the β-sheet-rich nitrophorin with a single histidine available for coordination (Figure 1). Other examples of single histidine coordinated hemes are horseradish peroxidase as well as the peptide-heme microperoxidases that will be discussed later.47−50 The second peptide in our set is HHHeme; the bis-histidine coordination is similar to neuroglobin, cytochrome b proteins (Figure 1),51 and a number of de novo designed, α-helical bundle peptides.52−56 Third, MHHeme was designed to offer a His-Met axial ligation similar to that found in cytochrome c. Finally, we employ AAHeme as an uncoordinating control peptide to monitor any background heme activity. These peptides produce a modest library that highlights the ability to tune heme-coordination and function within a peptide−amphiphile material through simple alterations in the primary coordination environment. Supramolecular Characterization. To test the hypothesis that supramolecular assembly has potential to effect the coordination environment surrounding the heme, it was crucial to analyze the morphology under various conditions. We find by transmission electron microscopy (TEM), atomic force microscopy (AFM), and dynamic light scattering (DLS) that the peptide−amphiphiles in HEPES buffer at pH 7 yield spherical micelles, Figure S1 (with the exception of AA). AFM shows that the spheres are ∼3−7 nm in diameter in both the presence and the absence of heme (at a ratio of 10 peptides:1 heme, Figures 2A,B and S1). Furthermore, DLS experiments suggest ∼7 nm diameter micelles in close agreement with the microscopy data (Figure S2). The spherical micelle formation is attributed to the large palmitoyl (c16) tail of the peptide− amphiphiles undergoing hydrophobic collapse in concert with electrostatic repulsion of the polar headgroup lysine residues. AAHeme was the only peptide to deviate from the spherical assembly where AFM, TEM, and DLS all showed fibers at neutral pH, Figure S1E. We presume that the lack of a bulky histidine residue at the aliphatic interior of the assembly eliminates steric repulsion, thus allowing the formation of fibers to occur. In 10 mM ammonium hydroxide at pH 10.5, the amine group on lysine is neutralized, thus decreasing the effect of electrostatic repulsion resulting in a micelle to fiber transformation (Figure 3)57 that was observed in all peptides studied as indicated by TEM and AFM (Figures 2C−D, S3, and S4). In addition, we employed HEPES at pH 10.5 (outside of the buffer range) to emphasize that HEPES, as a molecule, does not prevent the formation of fibers and that pH alone is adequate to

Figure 3. A cartoon representation of the spherical micelle formation in HEPES buffer at pH 7 and long aspect ratio nanofibers in 10 mM NH4OH at pH 10.5. 8499

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Table 1. Summation of Heme Coordination Characterization by Electronic Absorption Spectroscopy, Electron Paramagnetic Resonance Spectroscopy, and Binding Constant Analysis of the PAs, xyHeme, in Fiber (10 mM NH4OH, pH 10.5) and Micelle (HEPES, pH 7.0) Morphologiesa

a

EAS and EPR values of proteins with targeted coordination environments are listed for comparison. H.S. = high spin, L.S. = low spin.

therefore yields a spectrum comparable to that of free heme, Figure 4A, Table 1. Free heme in aqueous solutions readily forms aggregates yielding dramatically blue-shifted visible spectra from solubilized heme (Table 1).65,66 Furthermore, AAHeme yielded an entirely S = 5/2, high-spin EPR spectrum (Figure 4C) similar to that of free heme (Figure S8). In AAHeme, we suggest that heme aggregation is broken up and the molecule is solubilized (note: heme axial ligation is fulfilled by the coordination of a water molecule or hydroxide anion) but not coordinated by the peptide producing both the red shift in

the EA spectrum (in relationship to aggregated heme) and the signature high-spin spectrum from EPR, Table 1. Despite having one less histidine in its sequence than HHHeme, AHHeme micelles yield a bis-histidine heme coordination as the observed EPR spectrum is predominantly low-spin with g-values identical to those of HHHeme (Figure 4C). On the other hand, MHHeme micelles yielded a mixture of coordination states: a predominantly high spin, S = 5/2, EPR spectrum and an observable low-spin, bis-histidine contribution (Figure 4C). Consistent with the EPR data, the EA spectra for AHHeme and MHHeme micelles suggest a mixture of uncoordinated and bishistidine coordinated states (Figure 4A). This observation is consistent with the Soret and Q-band position and intensities, which represent averages between the spectra for bis-histidine heme coordination in HHHeme and embedded heme in AAHeme. The presence of bis-histidine axial ligation in AHHeme and MHHeme micelles is due to the greater degree of flexibility within the micelle assembly as compared to the rigid structure of the β-sheet fibers. Thus, bis-histidine coordination in micelles occurs with any peptide−amphiphile in our series that has a histidine, for example, xHHeme. Heme Coordinated to Fibers Characterization (pH 10.5). Next, we explore how a rigid supramolecular structure affects the heme-binding site. In fibers at high pH, the electronic absorption spectra indicate unique Soret and Q bands for each peptide, suggesting a variation in coordination environment from one sample to the next (Figure 4B and Table 1). Heme coordinated to HHHeme fibers exhibits a predominant signal attributed to bis-histidine coordination environment, similar to that of micelles, as indicated by the EA and EPR spectra (Figure 4B and D and Table 1). Note: Impurities of free heme are also detected, but the EAS is consistent with low-spin heme. Because AAHeme at neutral and high pH offers the same morphology, no significant spectroscopic changes were observed (Figure 4B and D and Table 1). We again suggest a lack of axial coordination to the peptide while the heme is fixed/solubilized in the matrix of the assembled fiber. Interestingly, MHHeme fibers yield spectra similar to those observed for AAHeme fibers, suggesting that heme is not coordinated but embedded in the peptide

Figure 4. Characterization of ferric heme coordination to different supramolecular constructs. Electronic absorption spectroscopy of heme (100 μM) bound to peptide (1 mM) at (A) pH 7.0 and (B) pH 10.5. EPR spectroscopy of heme (1 mM) bound to peptide (10 mM) at (C) pH 7.0 and (D) pH 10.5. Vertical lines and labels mark the high-spin (dashed lines) and low-spin (solid lines) states. AAHeme, blue; AHHeme, red; HHHeme, green; and MHHeme, purple. 8500

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then is available, the heme molecule will coordinate. The binding constant analysis determines which coordination site is the strongest, but most importantly highlights the significant influence the rigid secondary structure has over heme binding affinity. The EA spectra and EPR data found for heme coordination to the xyHeme series highlight a strong link between peptide sequence, molecular ordering, morphology, and heme coordination. HHHeme highlights our ability to design a peptide that maintains bis-histidine coordination when converting from micelles to fibers. MHHeme indicates that the bulky methionine side chain upon the ordering/formation of β-sheet rich fibers effectively blocks the histidine residues available for heme coordination in a micelle. Finally, AHHeme highlights a change in coordination environment concomitant with a morphological shift from (1) micelles yielding low-spin, bis-histidine coordination to (2) high density β-sheet containing fibers providing high-spin, single histidine axial ligation. These pHdependent changes serve as highly programmable features for the development of functional heme peptide materials. Electrochemistry. Redox behavior is another aspect where the binding site exerts control over heme in both natural and de novo proteins.32,50,52,53,70−72 Here, we demonstrate the ability to change the redox behavior of heme in our peptide materials through sequence and structure. In micelles, where the peptides exhibit similar coordination environments, AHHeme, HHHeme, and MHHeme exhibit similar midpoint potentials (EM) versus SHE: −315, −270, and −312 mV, respectively (Figure S10). When we analyzed the peptide fibers, the trend changed, such that AHHeme, HHHeme, and MHHeme all exhibit very different EM values versus SHE: −655, −333, and −442 mV, respectively (Figure S10). We attribute these results to the established variation in coordination environment between the micelle and β-sheet-rich fibrous morphologies. For example, there is negligible change in coordination state between HHHeme in micelles versus fibers, which extends to a minimal change in EM, but AHHeme yields a dramatic change that we attribute to different coordination modes, bis-histidine in micelles versus single-histidine in the β-sheet fibers. Gas Binding. Heme proteins often bind and transport small molecules like water, dioxygen (O2), or nitric oxide (NO) using coordination bonds. Each of these small molecules plays a vital role in signaling and catalysis, and when combined with the well-established properties of peptide amphiphiles produces multivariate functional materials that could be used in vasodilation (NO),73 neurotransmission (NO and CO),74,75 and O2 delivery or activation.76,77 We have chosen to explore carbon monoxide (CO) gas binding because CO serves as a redox inactive surrogate to these gases binding to ferrous (Fe2+) heme where (1) it offers major insight into small molecule accessibility of the xyHeme active site relevant to enzymatic activity and (2) provides information on how the peptide sequence can affect gas binding.78 The CO adduct was obtained by first chemically reducing the heme molecule in the peptide:heme assembly with a small amount of sodium dithionite in a nitrogen box. The sample then was transferred to a cuvette equipped with a rubber septum. Finally, the sample was sparged with CO for 30 s. UV/visible spectroscopy yields similar spectra when CO is coordinated to a histidine axially ligated ferrous heme, Figure S11. Therefore, infrared spectroscopy is employed because the heme binding pocket and ligand coordination directly influence the vibrational frequency of CO (νCO) either through enhanced backbonding or through ligand

assembly. This observation is further corroborated by the purely high-spin EPR spectrum and is in opposition to the mixture of high- and low-spin heme observed in MHHeme micelles. It should be noted that cytochrome c, which coordinates heme through histidine-methionine ligation, yields a low-spin, type I, highly anisotropic low-spin (HALS) spectrum with a g value of ∼3.3, further suggesting that we did not achieve the desired histidine-methionine coordination in MHHeme (Figure 1).67 We attribute the lack of heme coordination to the increased molecular ordering (i.e., β-sheets) within the assembly where the bulky methionine residue sterically blocks heme access to the histidine coordination site. Finally, heme coordinated to AHHeme fibers yields an EA spectrum indicative of coordinated heme but not typical of bishistidine axial ligation as it is blue-shifted to a value similar to that of nitrophorin, which possesses single histidine axial ligation (Figure 4B and Table 1). The EPR spectrum shows a predominantly high-spin species as expected for single histidine axially coordinated heme like nitrophorin.64,68 The value observed at gz = 3.71 is typical of a low-spin type I (HALS) spectrum and is observed in our control experiment where heme is analyzed in the presence of lysine (Figure S8).63 We conclude that the observed spectrum for heme coordinated to AH Heme fibers is predominantly a high-spin spectrum representative of a single histidine-coordinated heme with some propensity for unresolved low-spin states. Binding Constant Analysis. Peptide-to-heme stoichiometry and binding constants (Table 1) obtained from a series of titration experiments indicate a dependence on the flexible micelle structures when compared to the rigid fiber structures, Figure S9. We employed a binding analysis method typically used to analyze heme or transition metal binding to de novo designed proteins, eq S1.69 We modified the equation to yield a value n that represents the minimum number of peptides required to bind one heme. The results for micelles suggest a 3:1 (peptide:heme) stoichiometry, while fibers yield a 6:1 stoichiometry (Figure S9). Within the micelle construct, the lack of secondary structure yields a more flexible peptide environment, allowing for the more favorable bis-histidine coordination. As a result, the highest binding affinity (lowest Kd) is simply the peptide with more available histidines, HHHeme. MHHeme in micelles yields a less favorable bis-histidine coordination environment than AHHeme due to steric crowding from the bulky thioether at the heme-binding site. In contrast to the favorable bis-histidine coordination environment in micelles, the β-sheet-rich fibers offer a rigid structure that contributes significantly to heme coordination and binding affinity. While the HHHeme binding affinity is similar to that found for micelles, AHHeme binding affinity actually increases (lower Kd) despite the morphologically induced change to a single-histidine coordination environment. This increase in binding affinity is promoted by the rigid fiber structure and consequently the nonbulky alanine in the “distal” position of the heme-binding pocket. As a result, the opposite trend is observed in MHHeme, where the steric crowding of the bulky methionine that was observed in micelles is amplified by the rigid system and is reflected in the decrease in heme binding affinity (increased Kd). Despite the lack of a histidine containing coordination site, AA Heme does produce a “solubilization” curve, suggesting heme incorporation within the fiber construct. This helps to explain the overall mechanism of heme insertion such that the amphiphilic micelle or fiber first encapsulates the hydrophobic heme molecule. If a binding site 8501

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molecular construct immediately surrounding the heme molecule yielding a distribution of states. We focus our discussion on the major contributing peak. In the case of HHHeme, we assume one of the histidine ligands becomes uncoordinated, allowing CO binding to occur, yielding a distal histidine similar to that found in myoglobin. Again, the value obtained for HHHeme, νCO = 1961 cm−1, is higher than values obtained for myoglobin, but it is slightly lower than the value obtained for the micelle conformation. We suggest that the slight increase in heme−CO back-bonding within the fiber assembly is due to the presence of a distal histidine in a more rigid environment. When we produce a similar coordination environment to the distal-site mutants of myoglobin (e.g., His → Ala), we observe an increase in stretching frequency for AHHeme, νCO = 1971 cm−1, slightly greater than that found in the micelle structure for AHHeme. This is likely due to the lack of secondary structure in the micelles allowing uncoordinated histidine residues to interact with the heme−CO complex in the pocket, yielding a slightly lower νCO. As a result, the HHHeme CO frequency in fibers is 10 cm−1 lower than that found for AHHeme. We suggest that in the β-sheet enriched fibrous assemblies, CO serves as an excellent probe of the heme active site, indicating enhanced back-bonding due to a distal histidine in the case of HHHeme and reduced back-bonding due to the aliphatic alanine in the case of AHHeme. CO vibrational analysis for MHHeme is similar to that for AAHeme. This is consistent with the comparable electronic absorption spectra (Figure S11). While MHHeme was able to coordinate heme in micelles, the CO data are consistent with the lack of ferric heme binding to MH fibers. That is, the heme embeds itself in the fiber but without axial ligation to histidine. When comparing the CO vibrational spectra for AAHeme and MHHeme, a slight increase in the contribution of νCO = 1964 cm−1 is noticeable that may suggest some axial histidine coordination. The CO binding studies clearly highlight that the heme binding PAs are capable of binding gases like CO. Furthermore, detailed vibrational analysis highlights and supports our claims of controlling the coordination environment through careful sequence design and morphology. These results provide a basic understanding and some guiding principles for designing hemebinding peptides that can catalyze or transport and release gas on the basis of environmental triggers. Supramolecular Control of Enzymatic Activity. Many natural and synthetic heme proteins display peroxidase activity, a natural reaction that catalyzes the oxidation of high potential substrates using hydrogen peroxide (H2O2).31 We acknowledge the existence of other peptide:heme complexes, like microperoxidase (MP-11),79 de novo designed heme binding peptides,80−83 and even the demonstration of an active amyloid-β peptide,28 as well as heme solubilized micelles that yield peroxidase activity,84−88 but these systems do not probe the effects of supramolecular morphology. Here, we employed heme-peroxidase catalysis as a benchmark reaction to investigate specifically how supramolecular morphology in our PA−heme assemblies can control catalytic reactivity.89 We have chosen the established protocol where H2O2 activated by the xyHeme assemblies reacts with the colorless molecule 3,3′,5,5′tetramethylbenzidine (TMB) to yield the single electron oxidation product, TMB•+ (λmax = 652 nm) and water.90 All of these assays were carried out with 30 μM peptide, 1 μM heme in HEPES buffer at pH 7. Peptide assemblies were first prepared in either pH 7 HEPES (micelles) or 10.5 NH4OH

coordination enhancement/disruption as indicated in the wide range of observed stretching frequencies, νCO = 1949−1971 cm−1, Figure 5 and Table 2.78

Figure 5. Carbon monoxide vibrational analysis probing enzymatic capability and molecular structure. Infrared spectroscopy of heme−CO binding in the CO stretching frequency region at pH = 7 and pH = 10.5. AAHeme, blue; AHHeme, red; HHHeme, green; and MHHeme, purple. Hemin, 100 μM; peptide, 1 mM.

Table 2. Summary of CO Vibrational Analysisa pH = 7 peptide AAHeme AHHeme HHHeme MHHeme

νmaj (%) −1

1964 cm (57%) 1967 cm−1 (87%) 1963 cm−1 (96%) 1969 cm−1 (80%)

pH = 10.5 νmin (%) −1

1948 cm (43%) 1948 cm−1 (13%) 1974 cm−1 (4%) 1953 cm−1 (20%)

νmaj (%) −1

1965 cm (53%) 1971 cm−1 (58%) 1961 cm−1 (86%) 1964 cm−1 (63%)

νmin (%) 1948 cm−1 (47%) 1955 cm−1 (42%) 1977 cm−1 (14%) 1951 cm−1 (37%)

a

The major (νmaj) and minor (νmin) vibrational frequencies and their corresponding percent contributions are derived from Gaussian peak fitting (Figure S12).

For micelles at pH = 7, AHHeme, νCO = 1967 cm−1, HHHeme, νCO = 1963 cm−1, and MHHeme, νCO = 1969 cm−1 yield similar values (Figure 5, Table 2) due to their similar coordination environments consistent with EAS and EPR. For CO to bind in the micelles, one histidine must dissociate, resulting in a distalhistidine ligand. The observed vibrational frequencies are notably higher than those obtained for the analogous coordination environment of myoglobin at neutral pH78 (νCO = 1947 cm−1) likely because the micelle assembly lacks the more sophisticated structure of a fully folded protein. Mutations to various residues in the myoglobin active site result in an observed decrease in CO-heme backbonding, evidenced by an increase in stretching frequencies,78 νCO = 1965−1971 cm−1, consistent with the measured values here. In the case of AAHeme where fibers are formed regardless of environment and no discernible coordination to the peptide is observed, two vibrational states were found, νCO = 1948 and 1964 cm−1, with the former yielding the most intense peak, which is consistent with a five-coordinate heme−CO with no axial ligation to the peptide. Interestingly, with xyHeme fibers, which greatly influence the coordination environment of ferric heme, we observe different CO vibrational frequencies for each assembly (Figure 5). In general, the signals are broader or split when compared to the data at pH 7. This is most likely due to slight variations in the 8502

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Journal of the American Chemical Society Table 3. Michaelis−Menten Parameters for the c16-xyL3K3-CO2H Peptide Series Comparing Micelles and Fibersa

a

Free heme and MP-11 parameters are included for comparison. The values determined are from TMB oxidation with respect to varying the H2O2 concentration.

(fibers) with a 30-fold excess of peptide to heme (1.5 mM peptide, 50 μM heme). This 30:1 peptide:heme ratio (as opposed to the 10:1 peptide:heme used in material characterization) was employed to ensure that, upon a 50-fold dilution into HEPES, the heme would stay coordinated to the assembly. As a result, the observed peroxidase reactivity is generated by the heme bound to the assembly and not free heme, which displays peroxidase activity in the absence of peptide. The dilution process does not alter the peptide micelle and fiber structures (Figure S13), and heme remains coordinated (Figure S14). We also monitored free heme activity as well as microperoxidase to generate comparisons to an unprotected heme molecule and an optimized peroxidase. Michaelis−Menten kinetic analysis was performed to compare the peroxidase reactivity of micelles and fibers with respect to varying peroxide concentration (Figures S15 and 16), and the results are summarized in Table 3. We also measured the kinetics with respect to varying TMB concentration to indicate that our reported values are under saturated TMB conditions (Figure S17). Bleaching of the heme was not observed during the course of the experiments (Figure S18,19). The heme binding micelles, AHHeme and MHHeme, exhibit similar degrees of peroxidase activity, whereas their fibrous assemblies display a weak, baseline level of reactivity (Figure 6A,B). AHHeme was found to have the most dramatic difference between the two morphologies with the turnover number (kcat) being 16 times greater in micelles. Because of the relatively high KM in micelles versus fibers, the overall catalytic efficiency (kcat/KM) was found to be only 5 times greater in AHHeme micelles. We found a similar trend with MHHeme in which the turnover number (kcat) for micelles was 6 times greater than fibers. Turnover for HHHeme in micellar form was only 3 times greater than its fiber counterpart. In fact, when compared to the other peptides, HHHeme micelles exhibited a decrease in turnover while the fibers increased (Figures 6A,B and S16). This increase in kcat for HHHeme fibers may be attributed to a higher redox potential that is directly associated with its preferred low-spin, bis-histidine axial ligation. While AHHeme, HHHeme, and MHHeme micelles coordinate heme through bis-histidine axial ligation with comparable redox potentials, their difference in reaction rates may be attributed to the heme affinity. That is, HHHeme binds heme with the greatest affinity (Table 1) and shows the slowest rate of reaction among the micelles. For peroxidase activity to function properly, H2O2 must first react with the ferric heme before generating the active oxo-ferryl intermediate, compound I. If the bis-histidine coordination is strong enough, the reactivity would be slower because one of the histidines would have to dissociate prior to

Figure 6. Testing of peroxidase activity at pH 7 exhibits dependence on sequence and supramolecular morphology. Peroxidase activity, as seen by the oxidation of TMB, mediated by peptide−amphiphile series in either a (A) micelle or (B) fiber morphology. Heme activity with no peptide is represented by the dashed black line. Reaction conditions: peptide = 30 μM, heme = 1 μM, H2O2 = 6 mM, and TMB = 300 μM, HEPES buffer pH 7. (C) Representative Michaelis−Menten analysis for AHHeme, in micelles (solid red line) and fibers (dashed red line). Additional Michaelis−Menten analyses are available in Figure S15. (D) Total TMB oxidized upon completion of the peroxidase reaction. Solid bars represent micelles, and hashed bars represent fibers. Color code: blue, AA; red, AH; green, HH; purple, MH; and black, free heme.

reacting with H2O2. This binding affinity-based argument may hold for comparing the peptide micelles, but the fiber assemblies display a lower and baseline level of peroxidase reactivity with lower kcat and Km values. We therefore attribute the lowered activity specifically to the change in morphology from micelles to fibers, which, depending on the peptide sequence (e.g., AH), may involve a change in coordination environment that leads to a dramatic change in overall reactivity. For comparison, we measured the peroxidase activity of heme alone under our reaction conditions to highlight the effect of the peptide on peroxidase activity. Qualitatively, the peptide micelles enhance the catalytic ability of heme, while fibers decrease the activity (Figure 6A and B). Michaelis Menten kinetic analysis indicated that kcat/KM for free heme is 8503

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sickle cell disease, ischemia reperfusion, and malaria).30 When that micelle material reaches the target, this stimuli responsive material can react with the environment, triggering a change in morphology from micelles to fibers, all while sequestering and thus detoxifying heme by limiting its ability to generate reactive oxygen species. Furthermore, the well-established regenerative properties of PAs can be employed to provide a scaffold and stimulate the growth of healthy tissue.10

greater than that found for the peptide micelles, and kcat was lower suggesting that the heme molecule does not catalyze TMB over long periods of time and is evident in our analysis of total TMB oxidation (see below). This suggests that, while seemingly faster, the overall catalytic performance was weak. We also compared the peptide micelles with heme solubilized in surfactant micelles (heme in 1 wt % Tween 20, 10 nm micelles and 10 wt % SDS, 3 nm micelles) to highlight the peptides role in catalytic performance. Peroxidase activity of heme solubilized in surfactant micelles leads to slower TMB product formation than peptide micelles, Figure S20. In addition, inconsistent generation of both one- and two-electron TMB oxidation products in Tween and SDS micelles containing heme was not comparable to the results found for our peptide assemblies, suggesting that the heme coordinating ability of the peptide micelles plays an impactful role in generating controlled peroxidase activity. In contrast, peroxidase activity of microperoxidase-11 (an isolated heme-peptide segment from peroxidase) under our reaction conditions yielded a kcat/Km value ∼4000 times greater and a kcat value ∼500 times greater than the baseline activity of the fibers highlighting the significantly decreased catalytic ability of the fiber construct. We measured the final product concentration to compare the total amount of TMB oxidized (Figure 6D). We acidified our samples to yield the final two-electron oxidation product, TMBox, and measured the absorbance at 450 nm (ε = 59 000 M−1 cm−1). Our results are consistent with the kinetic data in that AHHeme and MHHeme were the best performers yielding 117 and 110 μM of TMBox, respectively. HHHeme produced 76 μM of TMBox. The fiber assemblies yielded a baseline amount of TMBox (10−25 μM) similar to that of our control example of heme (no peptide) in HEPES. HHHeme fibers represent an exception to this trend yielding 52 μM of TMBox. Consistent with the Michaelis−Menten kinetics, HHHeme fibers exhibit more activity than do the other fibrous assemblies. Therefore, our peptides demonstrate slow peroxidase activity as micelles, but when analyzed for peroxidase activity as fibers, they typically exhibit baseline reactivity similar to that found for our control sample with just heme. The most dramatic change in reactivity is within the AHHeme system in which the micelle heme peroxidase activity is enhanced, whereas the fibers significantly limit the reactivity most notably characterized by the 16-fold decrease in turnover number when comparing micelles to fibers. Many heme-containing proteins and peptide assemblies display peroxidase activity. In fact, myoglobin has been demonstrated to yield peroxidase activity,91 and, as mentioned earlier, cytochrome c in the presence of cancerous cells has been proposed to change from an electron transfer protein to a peroxidase.41 Here, we demonstrate a similar phenomenon in that, with the same peptide (AHHeme), we can modulate the reactivity just by changing the morphology from micelles to fibers, a potentially useful mechanism in a peptide amphiphile material. We acknowledge that our peptide micelles do not make great peroxidases, as indicated by our comparison with microperoxidase (i.e., the turnover frequency, kcat, and catalytic efficiency, kcat/KM, greatly exceed the values found for our peptide micelles). We suggest here that significantly limiting peroxidase reactivity of heme is a potential route to detoxify free heme in medicinal applications. In other words, we can imagine beginning with or delivering the apo-micelle to treat diseases that result in excess of free heme or hemolysis (e.g.,

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CONCLUSION We have clearly demonstrated the ability to control heme coordination and function through peptide sequence design and supramolecular structure. Morphological control is exhibited through changes in buffer choice and pH; for example, micelle and fiber assemblies can be formed. We have highlighted that different heme coordination environments with varying affinities are observed depending on the morphology and primary sequence/designed-binding site: AHHeme complexes where the micelle conformation yields low-spin, bishistidine ligation but the fiber yields high-spin, single-histidine coordination. With regard to eliciting function, we have highlighted the ability of the material to coordinate the redox inactive surrogate small molecule, carbon monoxide, which highlights both the active site accessibility of the heme and the ability of the heme to bind and potentially transport this molecule crucial to neurotransmission, and vasodilation. Finally, we have found a strong influence of supramolecular morphology on peroxidase activity where micelles exhibit enhanced catalytic activity over fibers composed of the same peptide that exhibit baseline catalytic activity. The catalytic activity in micelles can be further tuned through the primary sequence, with AHHeme and MHHeme displaying the highest catalytic efficiencies in the presented series. The function of the heme is crucial to producing advanced peptide materials and will be explored in future work where studies on more complex assemblies are underway. This peptide amphiphile system provides multiple avenues with which to control potential enzymatic activity that can ultimately be translated to the material’s functional properties. Sequence can be used to modulate the enzymatic rate, while gross structure can act as an on/off switch allowing for tuning of the reactivity as a function of environment. These results significantly impact molecular design strategies for functional peptide materials where we have discovered that supramolecular structure plays an essential role dictating heme function. For example, we will investigate fibrous structures that have potential use in anti-inflammatory signaling where the peptide assembly could be employed to sequester and break down toxic free-heme, resulting from sustained injuries (similar to the protein heme oxygenase I) while simultaneously exploiting known peptide amphiphile technologies that promote healthy tissue regeneration.92 Overall, we have demonstrated a new means of functionalizing current peptide−amphiphile technologies through the empirical design of peptides with engineered conformational changes that influence metal cofactor active sites resulting in controlled, protein-like, peptide materials.

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MATERIALS AND METHODS

Peptide Synthesis, Purification, and Characterization. The synthetic procedure for c16-AHL3K3-CO2H has been reported in our previous studies.24 The syntheses of c16-AAL3K3-CO2H, c16HHL3K3-CO2H, and c16-MHL3K3-CO2H, cleavage from the resin, 8504

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Journal of the American Chemical Society and RP-HPLC purification followed the same strategy as the previously reported peptide. MALDI-TOF MS (Bruker UltrafleXtreme MALDI-TOF) was used to identify the peptides; c16-AAL3K3-CO2H: calcd for C58H111N11O10 + [H+], 1122.85; found, 1122.933 m/z. c16AHL3K3-CO2H: calcd for C61H113N13O10 + [H+], 1188.87; found, 1189.006 m/z. c16-HHL3K3-CO2H: calcd for C64H115N15O10 + [H+], 1254.90; found, 1255.023 m/z. c16-MHL3K3-CO2H: calcd for C63H117N13O10S + [H+], 1247.88; found, 1248.864 m/z, Figure S21. Sample Preparation. Each peptide (3−4 mg) was dissolved in nanopure water (Millipore A10) to obtain a 1 wt % solution, c16AAL3K3-CO2H (1 wt %, 8.9 mM), c16-AHL3K3-CO2H (1 wt %, 8.4 mM), c16-HHL3K3-CO2H (1 wt %, 7.9 mM), and c16-MHL3K3CO2H (1 wt %, 8.0 mM). Hemin (Porcine, Sigma-Aldrich) was dissolved in DMSO (Sigma-Aldrich) to achieve a 10 mM stock solution. Note: Hemin/DMSO stock solutions were always made to ensure that the final DMSO concentration in the sample was less than 1% (v/v). Typically, 38 μL of a 1 wt % stock solution was dissolved in 260 μL of either HEPES (50 mM HEPES, 100 mM NaCl, pH 7.0) or 10 mM NH4OH, pH 10.5, to yield a 1 mM sample. The samples were then heated to 65 °C for 10 min and cooled back to room temperature to ensure formation of the supramolecular assembly. After the sample was cooled, 3 μL of the 10 mM hemin stock solution was added to the sample to yield 100 μM hemin. The samples were again heated to 65 °C for 10 min and cooled back to room temperature to ensure complete heme coordination. Titration experiments analyzed samples that contained 50 μM heme in either HEPES buffer or 10 mM NH4OH. Preassembled peptide was added to individual solutions containing heme such that the peptide concentration ranged from 0 to 1000 μM in 50 μM increments. The samples were equilibrated at room temperature for 1 h prior to UV/visible measurements. The experimental data were fit using a modified equation (see Supporting Information)69 to analyze for binding stoichiometry (n) as well as binding constant (Kd). Microscopy. Scanning electron micrographs were obtained with a JEOL 7500 field emission scanning electron microscope equipped with a transmission electron detector. Samples of varying concentrations were diluted 100-fold in water and drop cast onto a 400 mesh copper grid with a carbon support film (Ted Pella). After 1 min, the excess solution was wicked away and the sample was air-dried. Atomic force microscopy (AFM) images were obtained with a Veeco MultiMode 8 scanning probe microscope equipped with a silicon nitride tip for imaging soft-materials. The sample was prepared by drop casting 100 μL of a 200 μM (peptide) sample on freshly cleaved mica (Ted Pella) and allowed to incubate for 20 min. The excess sample was wicked away with filter paper, and the sample was dried prior to measurements. Secondary Structure Analysis. To analyze secondary structural formation in the absence of heme, circular dichroism spectroscopy (Jasco, Inc. J-815) was employed to analyze the typical n−π* transitions found for a β-sheet assembly. Samples were prepared by diluting the 1 mM peptide samples described in the previous section 5fold to yield a 200 μM sample. Additional secondary structural characterization was achieved with infrared spectroscopy (Thermo Scientific, Nicolet 6700 FT-IR spectrophotometer). Ten microliters of the samples described in the previous section were dropcast onto a 32 mm CaF2 plate (Sigma-Aldrich) and were air-dried. The thin films were aligned in the spectrophotometer, and the amide I vibrations in the region from 1500−1800 cm−1 were analyzed. Heme Coordination. Electronic absorption spectroscopy (Cary 50 UV spectrometers) was employed to monitor the key π−π* transitions typical of porphyrin derived molecules. The 1 mM peptide/ 0.1 mM hemin samples described earlier were transferred to a quartz cuvette with a 0.1 cm path length window (Starna Cells, Inc.) and analyzed from 300−800 nm. X-band continuous wave EPR experiments were carried out using a Bruker ELEXSYS E580 spectrometer operating in the X-band (9.4 GHz) and equipped with an Oxford CF935 helium flow cryostat with an ITC-5025 temperature controller. Samples for EPR (same preparation as described in the Sample Preparation) were concentrated to 1 mM hemin and 10 mM peptide with a 10 000 molecular

weight cutoff spin diafiltration system (EMD Millipore Inc., Amicon Ultra-0.5 Centrifugal Filter Unit with Ultracel-10 membrane). All experiments were performed at 10 K with modulation amplitude set to 1 and power set to 10 mW. Electrochemistry. The samples were placed in a spectroelectrochemical cell (1 mm quartz) equipped with a platinum mesh working electrode, platinum wire auxiliary electrode, and a Ag/AgCl reference electrode (Basi, Inc.). The samples were electrochemically reduced over a range from +200 to −700 mV versus SHE. Each applied voltage setting was allowed to equilibrate for a minimum of 20 min prior to UV/visible spectral acquisition (PerkinElmer, Lambda 950, UV/vis/NIR spectrophotometer). Midpoint potential analysis was achieved by fitting a standard Boltzmann curve to the obtained data (OriginPro 9.1). Carbon Monoxide Binding. The heme ferrous state was obtained through chemical reduction by adding 5 μL of a concentrated sodium dithionite (Sigma-Aldrich) solution (100 mg/mL) into a preassembled 1000 μM peptide/100 μM hemin solution (300 μL) in an eppendorf tube. All samples were equilibrated and handled in an inert, nitrogen atmosphere (Plas Laboratories Inc. 830 Series Compact Glove Box). Carbon monoxide (99.99%, Airgas) was added directly through the solution in the eppendorf tube for 30 s. Ten microliters of the solution was dropcast onto a CaF2 plate where CO(g) was gently blown over the droplet resulting in a thin film of the PAHeme material. The samples were than analyzed by FTIR spectroscopy (Thermo Scientific, Nicolet 6700 FT-IR spectrophotometer). The samples were stable against oxidation during the course of the experiments. All obtained data were fit to a double Gaussian peak distribution due to the pronounced shoulders in some of the spectra. Peroxidase Activity Assay. Peroxidase activity was monitored using a Varian Cary 50 spectrophotometer and the kinetics software package. All kinetics were performed in a solution of HEPES buffer, pH 7 (50 mM HEPES 100 mM NaCl). The following stock solutions were used: TMB (10 mg/mL, 41.6 mM in DMSO), hydrogen peroxide (100 mM diluted in HEPES); H2O2 stock concentration was standardized by the method of Klassen et al.93 and by UV/vis (ε230 = 72.8 M−1 cm−1). Stock solutions of 50 μM heme and 1500 μM PA in HEPES (micelles) or 20 mM NH4OH (fibers) were equilibrated overnight in the dark. In disposable plastic cuvettes, 2 mL of HEPES buffer was added. TMB was added at a concentration of 300 μM. H2O2 was varied from 1.0 to 10 mM. To initiate the reaction, 40 μL of the heme stock solution was added and mixed via pipetting to yield 1 μM heme and 30 μM PA. The generation of the single electron oxidation product of TMB was monitored (ε652 = 39 000 M−1 cm−1) over a 5 min period collecting data points every 10 s. Initial velocities (vo, μM s−1) were determined by fitting the linear region of the kinetics (Figure S14). The initial velocity was then plotted versus peroxide concentration (Figure S15). Michaelis−Menten curves were fit using Origin 9.1 software to the equation (v0 = kCat[E]0[S]/(KM + [S])). The total amount of oxidized TMB was calculated after allowing the reaction to go to completion for 1 h. The reaction then was diluted 10-fold in 1 M HCl to yield the final, yellow TMB oxidation product. The absorption was monitored at 450 nm to determine the total amount of TMB oxidized, ε450 = 59 000 M−1 cm−1. Microperoxidase11 (Sigma-Aldrich) and free heme were prepared at 50 μM in HEPES. SDS (10 wt % in HEPES, pH 7, diameter = 3 nm) and TWEEN 20 (1 wt % in HEPES, pH 7, diameter = 10 nm) micelles were characterized by DLS (data not shown) to confirm correct micelle diameter. Heme was added to a final concentration of 50 μM. The control solutions were aged overnight in the dark to be consistent with the sample preparation of our peptides. Because of its high degree of reactivity, 10 μL of the MP-11 stock (250 nM final concentration) was added to initiate the reaction of the control experiment.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/jacs.7b01588. 8505

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AFM, DLS, SEM, FTIR, CD, EPR, binding constant analysis, electrochemistry, CO binding, dilution stability, Michaelis Menten plots, heme bleaching studies, and mass spectroscopy analysis including Figures S1−S21 and eq S1 (PDF)

AUTHOR INFORMATION

Corresponding Author

*[email protected] ORCID

Lee A. Solomon: 0000-0003-1471-9510 H. Christopher Fry: 0000-0001-8343-5189 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We wish to thank Tijana Rajh for helpful discussions related to the manuscript in addition to her assistance with EPR. J.K. participated in this work through the Illinois Mathematics and Science Academy’s Student Inquiry and Research (SIR) program supported in part through the State of Illinois Education Assistance Fund. This work was performed at the Center for Nanoscale Materials, a U.S. Department of Energy Office of Science User Facility under contract no. DE-AC0206CH11357.

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DOI: 10.1021/jacs.7b01588 J. Am. Chem. Soc. 2017, 139, 8497−8507

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DOI: 10.1021/jacs.7b01588 J. Am. Chem. Soc. 2017, 139, 8497−8507